Diagnosis of Canine Heartworm Infection

Margaret C. Barr, DVM, PhD; Elizabeth P. Boynton, DVM; Peggy L. Schmidt, DVM, MS, Diplomate ACVPM; Frank Bossong, DVM; and Gary R. Johnston, DVM, MS, Diplomate ACVR

Factors influencing the spread of canine heartworm (CHW) include:

• Increased movement of domestic dogs across the U.S.
• Fewer mosquito and vector control programs in response to concerns about pesticide safety
• Lack of compliance with prevention programs
• Increased infection of nondomestic canid reservoir hosts, such as the American coyote.²

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Editor’s Note: This article was originally published in July 2011. Please use this content for reference or educational purposes, but note that it is not being actively vetted after publication. For the most recent peer-reviewed content, see our issue archive.

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Heartworm Hosts

While dogs are considered to be definitive hosts for heartworms, D immitis may infect many different species, including indigenous canids, such as wolves, foxes, and coyotes; many species of felids; and occasional incidental hosts, such as deer, ferrets, raccoons, horses, and primates.

The noncanid species are typically dead-end hosts; however, even a single heartworm in some animals may cause significant pulmonary disease, especially in cats.^2,3 In humans, dying adult worms are associated with the development of pulmonary granulomas, which may be misdiagnosed as lung cancer on thoracic radiographs.³

DIAGNOSIS OF INFECTION

In-clinic diagnosis of CHW infection is typically accomplished through detection of circulating microfilaria or soluble CHW antigen, and is dependent upon allowing adequate time from potential infection date to test date to account for the prepatent period (see Life Cycle of Heartworms). Because of this lag time, a general rule of thumb is to wait until a dog is 7 months of age before testing for the first time.⁴

Life Cycle of Heartworms

1. The life cycle of a patent CHW infection starts with introduction of L3 larvae from an infected mosquito into the canid host.

2. In the dog, the larvae mature into L4 larvae and then into adult worms that localize in the pulmonary arteries and occasionally in the right cardiac ventricle.

3. Following a prepatent period of 6 to 7 months, microfilariae are produced via sexual reproduction and are released into the dog’s bloodstream, where they may remain viable for months to years.

4. Mosquitoes feeding on the infected host ingest the microfilaria, which then migrate to the Malpighian tubules where they mature into first-stage larvae.

5. Ultimately, the larvae mature to the L3 stage in the salivary glands of the mosquito and become infective, thus completing the heartworm life cycle.

Courtesy of the American Heartworm Society ©2007

When to Test

Periodic testing, usually yearly, is recommended for all dogs, and it is advisable to retest whenever changing chemoprophylaxis agents or starting prophylaxis after a period of noncompliance. When changing heartworm preventive medication, additional testing 4 months after switching products is required to evaluate the efficacy of the original product. This process is especially important in light of recent suspicions of growing CHW resistance to macrocyclic lactones in some regions of the U.S. (see Prophylaxis Update).

Prophylaxis Update

Chemoprophylaxis is the primary weapon available to prevent heartworm infections in individual animals and controls spread of this parasite by reducing infections in the primary reservoir population. Macrocyclic lactones (ivermectin, moxydectin, selamectin, and milbemycin oxime) are the most commonly used heartworm preventives. The American Heartworm Society’s current guidelines4 for control of CHW infections in dogs include:

• Begin CHW prophylaxis in puppies as early as possible, usually before 8 weeks of age.
• Year-round chemoprophylaxis (even in areas where cold weather interrupts the CHW life cycle) is highly recommended to increase compliance.
• Although collies and some other dogs are sensitive to macrocyclic lactones, commercially available products for dogs used at the recommended doses are considered to be safe in all breeds.
• Reports of lack of efficacy of macrocyclic lactones in some areas of the U.S. are being investigated; currently, there is no strong evidence of widespread emergence of resistant strains of CHW. Lack of treatment and lack of compliance are of much greater concern.

The issue of apparent lack of efficacy is yet to be resolved. A recent re- port detailed genetic changes in some populations of heartworms that may confer greater resistance to chemoprophylactic agents.⁵ Whether or not these changes translate to the emergence of macrocyclic lactone-resistant strains of heartworms in endemic populations remains to be seen.

Microfilaria Detection

Microfilaria may be visualized in whole blood using a direct smear technique or from blood concentrated using a modified Knott’s test or membrane filtration. Compared to other methods of heartworm detection, blood smears and concentration techniques are relatively insensitive; therefore, a test should not be considered negative until at least 1 mL of blood has been examined.⁴ However, the sensitivity of the modified Knott’s test is greater than that of the direct smear method when samples contain < 50 microfilaria per 1 mL of whole blood6 and the test provides enhanced ability to distinguish D immitis from other canine microfilaria.

Microfilaria tests are now viewed as supplemental assays, secondary to the more sensitive heartworm antigen tests. Microfilaria detection can be used to:

•  Validate a positive antigen test
•  Determine if the dog is a potential reservoir for infection
•  Identify dogs at risk for severe reaction to treatment with microfilaricides.

Limitations of microfilaria tests include:

•  The fact that about 20% of heartworm-infected dogs are not microfilaremic (occult infection), usually because only 1 sex of worm is present or the infection is prepatent.⁴
•  The use of macrolide preventives may result in clearance of microfilaria from the blood of treated dogs after several months of use.^2,4

Antigen Detection

Enzyme-linked immunosorbent assay (ELISA) technology (Figure 1) was used in the first in-clinic CHW antigen tests; however, several current in-clinic CHW antigen tests are immunochromatographic (ICT) assays (Figure 2).

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Figure 1. ELISA Test Procedure

1. Anti-CHW antibodies (Ab) linked to an enzyme are mixed with whole blood, plasma, or serum, allowing them to bind to heartworm antigens (Ag) in the test sample.

2. The conjugate–sample mixture is then applied to the sample port (S) and allowed to flow across a membrane on which spots of additional anti-CHW Abs have been bound to detect low (L) and high (H) antigen levels.

3. A wash solution is used to remove unbound enzyme conjugates. The wash is followed by a substrate solution that acts with the bound enzyme to produce a color change, indicating a positive test.

4. Positive (C) and negative (N) control dots are also included on the membrane. Presence of a color change in the positive control is required for a valid test, and sample dots must also have stronger color development than negative control.

5. When no positive control dot develops or the negative control has high reactivity, the test is considered invalid and must be repeated.

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Figure 2. ICT (Lateral Flow Test) Procedure

1. A drop of the sample (whole blood, serum, or plasma) is added to the sample port (S) of the test device.
2. The sample drop reconstitutes the conjugate of gold particles linked to anti-CHW antibodies (Ab) dried on a pad in the test device. A chase buffer is often used to improve flow.
3. The CHW antigen (Ag) in the sample and the reconstituted conjugate flow down the test strip over 2 specific areas.

•  The first area is a line containing anti-CHW Ab (test line, T), which is immobilized on the strip. If CHW Ag is present, the anti-CHW Ab will capture it along with bound particle conjugate, causing a visible line to form.
•  Excess conjugate migrates past this line and over the control (C) area, where conjugate-specific antibodies capture the particle conjugate causing a visible line to form.

Positive samples must produce obvious lines in both the test and control sections of the strip for the test to be valid. Negative samples produce an obvious line only in the control section. Invalid tests, indicated by a line in the test area but not the control area, or by no lines in either area, must be repeated.

Even though they are more sensitive than microfilaria detection, CHW antigen tests:

• May give false negative results in animals that are infected only with male worms or with very few female worms; antigen testing fails in this case because the monoclonal antibodies used in the tests primarily detect female-specific antigens⁷
• Are not always precise with regard to correlation between the amount of heartworm antigen in the blood and the number of adult female worms.

Commercial CHW antigen tests evaluated during 2001 to 2002 were found to reliably detect at least 84% of CHW infections in which 3 or more female adult worms were present, but sensitivity decreased when fewer than 2 adult females were present.^8,9

DNA Amplification

In addition to detection of microfilaria and/or heartworm antigen, other tests may be useful in diagnosing CHW infection, especially in microfilaria-negative dogs or dogs with low worm burdens. Recently, polymerase chain reaction (PCR) technology has been used for the detection and identification of specific microfilaria in peripheral blood samples.

• Amplification of DNA from circulating microfilaria in a multiplex PCR assay gave reliable positive results for samples containing as few as 4 microfilaria per mL and distinguished D immitis infections from D repens infections.¹⁰
• Another multiplex PCR assay was used to distinguish between 6 different species of microfilariae in dogs and resolve discrepancies in morphologic diagnosis by 2 diagnostic laboratories.¹¹ Molecular speciation identified microfilaria from several samples as D immitis, differing from an original morphologic diagnosis as Acanthocheilonema (Dipetalonema) reconditum.

Currently, the clinical relevance of PCR testing for CHW is low because it is not available through large commercial laboratories.

Imaging

Ultrasound and radiology can be useful tools for diagnosing and staging CHW disease. Radiographic findings in a heartworm positive dog can be variable due to several factors such as:

•  Length of infection and corresponding disease progression
•  Size of the population of worms present
•  Worm location in the cardiopulmonary system.¹²

In an animal with recent CHW infection, there may be no radiographic changes present; however, as the duration of infection increases, common changes include (Figure 3):

•  Right ventricular enlargement
•  Pulmonary vasculature dilation
•  Parenchymal lung infiltrates.

Figure 3. Ventrodorsal thoracic radiograph from a 6-year-old neutered male pug that was presented for labored breathing. The patient spent the last 3 years in Louisiana and had not been receiving heartworm preventive. The ventrodorsal thoracic radiograph revealed blunted, enlarged, and tortuous pulmonary arteries (large arrows) and an enlarged pulmonary trunk (small arrows). These radiographic findings are consistent with heartworm disease.

In dogs with severe disease, the pulmonary arteries can appear tortuous or have nodular opacities. Hepatomegaly and pleural effusion may also be seen radiographically if the progression of the CHW infection leads to right-sided heart failure.¹²

On ultrasound, adult worms can appear as parallel hyperechoic lines present in the pulmonary artery and occasionally in the right ventricle (Figure 4). Echocardiography can also be a useful tool in detecting the extent of right heart enlargement and functional impairment due to CHW infection.4,13 The presence of worms in the orifice of the tricuspid valve in dogs with hemoglobinuria is indicative of caval syndrome.⁴

Figure 4. B-mode echocardiogram obtained in the left parasternal approach during diastole. The right ventricular wall (RVWd), right ventricle (RVd), and interventricular septum (IVSd) are identified. The heartworms (arrow) are identified in the right atrium (RAd).

EVALUATION OF CHW ANTIGEN TESTS

Given that in-clinic antigen assays are now the primary method for detecting CHW infection in dogs, it is important to know how well current tests are performing. In the past 10 years, some tests have been reformulated and new tests have been introduced.

We evaluated the performance of 4 immunoassays currently available for point-of-care diagnosis of CHW infection. Samples from a total of 114 dogs were tested, including 51 negative and 10 positive ethylenediaminetetraacetic acid (EDTA) whole blood samples, and 15 negative and 37 positive serum or plasma samples (Table 1).

Test Results

Tests were designated A, B, C, or D. Concordance among the tests was excellent.

•  The only discordant test was a weak positive on Test A that was negative on all other tests. The sample, which was whole blood from a healthy, low-risk dog, was rerun on all tests with the same results using both whole blood and plasma.
•  None of the 114 samples run on Tests A, C, and D were invalid; 2 samples run on Test B did not develop a positive control line and were declared invalid. The same samples were repeated successfully on additional Test B units.

The ability of the CHW tests to correctly identify CHW diseased dogs (sensitivity) and CHW disease-free dogs (specificity) was determined using the described criteria for identification of diseased and nondiseased samples (Table 2). Tests B, C, and D each demonstrated 100% sensitivity and 100% specificity, while Test A demonstrated 100% sensitivity and 98.5% specificity.

Potential Study Bias

Some potential for bias existed in the performance of diagnostic tests in this study.

•  Heartworm infected and noninfected animals were selected from distinctly different populations rather than from a spectrum of diseased and nondiseased subjects.¹⁴
•  CHW infected animals were all artificially infected. Of these 47 dogs, we received information on the infection dose for 25 dogs, with necropsy data available on 6 dogs.
•  The worm burdens in the artificially infected dogs were similar to the mean number of worms reported for naturally infected dogs in Michigan, higher than those reported for Maine, and lower than those reported for Florida.^8,15 However, marginally infected animals (worm burden of < 2 adult females), which can represent the largest numbers of false negative animals, were underrepresented and, therefore, our reported sensitivity values may be artificially elevated.

On the other hand, reported specificity values may be artificially elevated due to incorporation bias within the CHW negative group (see Standards for Heartworm Testing).¹⁴ The accepted gold standard is absence of adult worms at necropsy, but our disease-free population was defined by alternative criteria that included previous negative CHW antigen test results. This can lead to significant bias and artificial elevation of test specificity.

Minimizing Bias

The likelihood of bias was minimized through selection of traits that reduced the likelihood of these subjects to be infected including:

•  Disease-free clinical presentation (all dogs)
•  Residence in a low-risk area (all dogs)
•  History of administration of CHW preventative (most dogs)
•  Previous negative test results (multiple times in some dogs).

However, because the previous negative test results were determined by 1 of the 4 test kits (Test D) utilized in this study, the potential for bias, although small, is still present.

CONCLUSION

In most cases, diagnosis of CHW infection is relatively straightforward and easily accomplished through in-clinic testing. However, an understanding of the heartworm life cycle and recognition of the limitations of current diagnostic procedures are critical to successful diagnosis.

Two major limitations of most CHW tests are:

•  Inability to detect infection during the long prepatent period
•  Decreased sensitivity of detection when very few or no female worms are present.

Commercial CHW antigen tests have high sensitivity and specificity but performance may be compromised when these limitations are encountered.

Acknowledgements

We gratefully acknowledge veterinary technicians Kathryn Gresco, Eva Jaeger, and Kimberly Holt for their help with the CHW screening clinic. We also thank Dr. Brenda Knowlton and student representative Bridget Morton of Pfizer Animal Health for helping plan and organize the clinic.

CHW = canine heartworm;

EDTA = ethylenediaminetetraacetic acid;

ELISA = enzyme-linked immunosorbent assay;

ICT = immunochromatographic test;

PCR = polymerase chain reaction

Disclosure Statement

Partial funding for this study was provided by SafePath Laboratories (safepath.com).

Margaret C. Barr, DVM, PhD, is a professor of virology and immunology and serves as a coleader and facilitator for the Veterinary Basic and Clinical Sciences problem-based learning (PBL) course at the Western University of Health Sciences College of Veterinary Medicine. Dr. Barr is also the project director for the Snow Leopard Functional Genomics Initiative and serves on the board of directors for Pegasus Rising, an organization that provides equine-assisted therapy to veterans. Her interests include translational research on diagnostic assays for infectious diseases, nontraditional methods of immunization, emerging and reemerging infectious diseases, and host immune responses to infectious diseases. Current research projects include investigation of the molecular epidemiology of canine parvovirus and rickettsial agents in southern California. Dr. Barr received her DVM from Auburn University and her PhD from Cornell University.

Elizabeth P. Boynton, DVM, is the director of the Hill’s Wellness Center, directs the professional communication curriculum, and facilitates problem-based learning and clinical skills laboratories at Western University of Health Sciences College of Veterinary Medicine. She is a member of the California Veterinary Medical Association’s House of Delegates and House Advisory Committee. Prior to her position at WesternU, Dr. Boynton worked in private practice and then filled a new position as community practice veterinarian at Univeristy of Minnesota Veterinary Teaching Hospital. She was presented with the Faculty Appreciation Award twice as well as the Minnesota Veterinary Medical Association’s Outstanding Faculty Award. She received her DVM from University of Minnesota.

Peggy Schmidt, DVM, MS, Diplomate ACVPM, is an associate professor of population health and epidemiology at Western University of Health Sciences College of Veterinary Medicine. Dr. Schmidt serves as director of College Outcomes Assessment and coordinator of the WesternU DVM/MPH dual degree program. Her areas of research interest include evidence-based veterinary medicine, veterinary public health, infectious diseases, and veterinary education. Dr. Schmidt received her DVM from University of Minnesota. After working in a mixed-animal practice in western Minnesota, she joined Iowa State University College of Veterinary Medicine as an instructor while pursuing an MS degree in veterinary preventive medicine.

Frank Bossong, DVM, is an assistant professor in shelter medicine at Western University of Health Sciences College of Veterinary Medicine. He is also the head clinician for the Veterinary Ambulatory Community Service (VACS), a second year problem-based learning facilitator, and a course leader for the third-year internal medicine course. Dr. Bossong received his DVM from University of Georgia. After working as an associate for 4 years at an AAHA-accredited hospital in Pasadena, California, Dr. Bossong pursued his interest in shelter medicine. He established an on-site veterinary clinic, medical policies and procedures, and a fourth-year veterinary rotation at the San Gabriel Valley Humane Society, where he currently serves on the board of directors.

Gary R. Johnston, DVM, MS, Diplomate ACVR, is a professor of radiology at Western University of Health Sciences College of Veterinary Medicine and teaches all levels of the curriculum, facilitates problem-based learning, and participates in clinical research. Dr. Johnston received his DVM from Washington State University and then completed a rotating small animal medicine internship, MS, and residency in radiology at University of Minnesota. He remained at the University of Minnesota as a radiologist, progressing from assistant professor to full professor. Dr. Johnston spent 3 years at Washington State University College of Veterinary Medicine before joining the WesternU faculty.